REGULATION OF HEPATIC GLUCONEOGENESIS …hypoxia (with/without yS-adrenoreceptor blockade) on tissue glucose and glycogen levels, and on the kinetic behavior of two key enzymes involved

t -xp. Biol. 147, 169-188 (1989) 169

nted in Great Britain © The Company of Biologists Limited 1989

REGULATION OF HEPATIC GLUCONEOGENESIS ANDGLYCOGENOLYSIS BY CATECHOLAMINES IN RAINBOW

TROUT DURING ENVIRONMENTAL HYPOXIA

BY PATRICIA A. WRIGHT*, STEVE F. PERRY AND THOMAS W. MOON

Department of Biology, University of Ottawa, Ottawa, Ontario,Canada KIN 6N5

Accepted 28 June 1989

Summary

This study tests the hypothesis that catecholamines regulate glucose availabilityduring hypoxia in the rainbow trout by activating glycogen phosphorylase (GPase)while inhibiting pyruvate kinase (PK) in the liver. The net result would be anincrease in liver glycogenolysis and a reduction of glycolysis and/or enhancementof gluconeogenesis. We used the criteria of Stalmans & Hers (1975) and reportmuch lower resting percent GPase a (active) values (20-30%) than thosepreviously published. Dorsal aortic injections of epinephrine or norepinephrineincreased plasma glucose (16-46%), had no effect on liver or muscle glycogenlevels, decreased the activity of PK, and increased total and percent GPase aactivities. Pre-treatment with the /3-adrenoreceptor antagonist propranolol elimi-nated these effects. During moderate hypoxia, plasma glucose remainedunchanged, while lactate levels increased fourfold. When fish were pre-treatedwith propranolol, hypoxia depressed plasma glucose levels (—26%), total andpercent GPase a, and increased PK activity, suggesting that hypoxia mediated thedephosphorylation of these enzymes. We conclude that catecholamines stimulatehepatic /3-adrenoreceptors during hypoxia and sustain plasma glucose levels bynullifying the deleterious effects of hypoxia on metabolic function. The specificmetabolic consequences of these catecholamine-mediated effects are an increasein the activity of the active form of GPase and a reduction in PK activity, whichsuggests an activation of glycogenolysis and an inhibition of glycolysis and/oractivation of gluconeogenesis, respectively.

Introduction

When trout encounter environmental hypoxia they respond initially by loweringmetabolic rate, then by increasing anaerobic metabolism (Boutilier et al. 1988).

•Present address: Laboratory of Kidney and Electrolyte Metabolism, National Institutes ofalth, Building 10, Room 6N307, Bethesda, MD 20892, USA.

y words: glucose, hypoxia, catecholamines, trout, liver.

170 P. A . WRIGHT, S. F. PERRY AND T. W. M O O N

Some tissues are glucose-dependent (e.g. nervous tissue), but have little reservecapacity and, for this reason, it is important that plasma glucose levels areregulated. Moreover, rapid increases in glucose availability are often required tomeet the increased metabolic demands of tissues. It is not surprising, therefore,that catecholamine levels, well known as mediators of rapid physiological change,increase markedly in arterial blood during exposure to low environmental oxygen(Butler et al. 1979; Tetens & Christensen, 1987; Fievet et al. 1987; Boutilier et al.1988). The purpose of this study was to determine whether catecholamines play arole in the regulation of glucose metabolism in vivo during short-term moderatehypoxia.

Catecholamines exert hyperglycemic effects in fish (e.g. Thorpe & Ince, 1974;deRoos & deRoos, 1978; Morata et al. 1982; Ottolenghi et al. 1984, 1985, 1986;Janssens & Lowrey, 1987; Mommsen et al. 1988), which are due, in part, toactivation of glycogenolysis (Nakano & Tomlinson, 1967; Birnbaum et al. 1976;Rush & Umminger, 1978; Ristori & Laurent, 1985; Mommsen et al. 1988).Epinephrine also activates gluconeogenesis, but its relative importance is notclear. Mommsen et al. (1988) reported that glycogenolysis accounted for 97 % ofglucose production in trout hepatocytes with or without epinephrine. Ottolenghiet al. (1985), however, concluded that in isolated, perfused catfish livers, glyco-genolysis dominated at high doses of epinephrine, but gluconeogenesis was thedominant source of glucose at low doses.

The role of catecholamines in glucose metabolism has often been studiedin vitro, but less frequently in intact fish. One of the difficulties with usinghepatocytes in vitro is that freshly isolated cells are in negative glycogen balanceand rates are unlikely to reflect those found in situ (Mommsen et al. 1988). In thisstudy we use intact fish to eliminate this in vitro preparation artifact.

A second problem addressed by this study relates to methodological differencesin the measurement of glycogen phosphorylase (GPase) activity. Wosilait &Sutherland (1956) first separated the two forms of GPase (b, inactive; a, active) bymeasuring activities in the absence and in the presence of AMP, respectively.Unless caffeine is present, however, the activity of rat liver GPase b is significantlystimulated by AMP (Stalmans & Hers, 1975). This may explain the consistentlyhigh liver resting percent GPase a [= a/(a+b)] values reported in the literature(90-95%, Ottolenghi etal. 1985, 1986; 83%, Perry et al. 1988; 72%, Mommsenet al. 1988). Such high percent GPase a values suggest that GPase is almostcompletely activated at rest, a situation which affords little possibility of regulatorycontrol. In the present study, we rectified this technical problem by applying thecriteria of Stalmans & Hers (1975) to GPase activities.

We have analyzed the individual effects of epinephrine, norepinephrine andhypoxia (with/without yS-adrenoreceptor blockade) on tissue glucose and glycogenlevels, and on the kinetic behavior of two key enzymes involved at critical points inglucose metabolism, glycogen phosphorylase and pyruvate kinase. PK wasmeasured because of its key role in regulating metabolic flux through ^ Jand gluconeogenesis (Pilkis et al. 1988).

Glucose regulation during hypoxia in trout 111

Materials and methodsAnimals

Rainbow trout (Salmo gairdneri) of both sexes, weighing 150-250 g, wereobtained from Thistle Springs Trout Farm (Ashton, Ontario) and maintained asreported previously (Perry & Vermette, 1987) at seasonal temperatures(15-18°C). Preliminary PK kinetic properties were detennined on resting intactfish, held in individual, opaque, Perspex chambers. Fish were killed by a sharpblow to the head, the liver was immediately removed, and frozen betweenaluminum blocks cooled in liquid N2. Tissue samples were stored at — 70 °C untilanalysis.

In all other experiments, trout were anesthetized, surgically fitted with a dorsalaortic (DA) catheter (Soivio et al. 1972), and left to recover for 48 h in individualPerspex chambers. Three series of experiments were performed (series I, II andIII) and in each series there were four separate groups of fish (Table 1).Experiments were performed over the summer season when ambient watertemperature increased from 15 °C to a peak of 18°C. Fish in series I were held at15 °C, series II at 18°C and series III at 16°C.

Experimental protocol

In each series, fish were pre-injected with either heparinized, physiologicalsaline (Wolf, 1963) or propranolol, which was followed 1 h later by the experimen-

Table 1. Experimental design of series I, II and III

Series IEpinephrine

Series IINorepinephrine

Series IIIHypoxia

Experiment

Saline controlEpinephrine

Propranolol controlPropranolol +epinephrine

Saline controlNorepinephrine

Propranolol controlPropranolol +norepinephrine

Saline controlHypoxia

PropranololPropranolol +

hypoxia

Pre-injection

SalineSaline

PropranololPropranolol

SalineSaline


SalineSaline


Experimentalinjection/treatment

SalineEpinephrine

SalineEpinephrine

SalineNorepinephrine

SalineNorepinephrine

Normal waterHypoxic waterNormal waterHypoxic water

172 P . A . WRIGHT, S. F. PERRY AND T. W. M O O N

tal treatment of saline, epinephrine or norepinephrine injection, or exposure toexternal hypoxia (Table 1). Propranolol was dissolved in heparinized salineimmediately before use, adjusted to pH 7-8, and injected over a 30 s period into theDA catheter. The volume injected was 0-2 ml at a dose of 2 mg kg"1 body mass andwas followed by an infusion of 0-3 ml of saline to flush the catheter. The calculatedcirculating level of propranolol was approximately 5xlO~5moll"1, about threeorders of magnitude greater than peak catecholamine levels (see Fig. 2). Epineph-rine and norepinephrine solutions (bitartrate salts; pH7-8) were preparedimmediately prior to infusion. The volume injected was 0-2 ml at an appropriatedose to achieve a circulating epinephrine/norepinephrine concentration of5xl0~8moll~1, followed by a 0-3 ml saline bolus to flush the catheter.

In series I and II, an initial blood sample (0-8-1-0 ml) was collected 30min afterthe pre-injection and a final blood sample (0-8-l-0ml) was collected 15min afterthe experimental injection (Table 1). The initial blood sample was taken forcontrol measurements, and compared to blood samples taken 15min after theexperimental injection.

In series III, the degree and duration of hypoxia were chosen to induce amarked increase in circulating catecholamines within a minimum time (Boutilieret al. 1988). External hypoxia was achieved by gassing the inflowing water with100% N2 in a counter-current gas-exchange column. By carefully adjusting thewater and gas flow rates through the column, the oxygen tension of the inflowingwater (PwO2 = 7-l ±0-4kPa, S.E.) was kept constant for a 30min experimentalperiod.

In all series, fish were killed by a sharp blow to the head immediately after thefinal blood sample had been withdrawn through the DA catheter. The liver wasexcised and frozen between aluminum blocks cooled in liquid N2. White musclesamples (0-5-1-0 g) were then excised and quickly frozen in the same manner.Tissue samples were stored at —70°C until analysed (<24h for glucose andglycogen, <4 days for liver enzyme analyses).

Analytical procedures

Blood was analysed for pH (pHe), red cell pH (pHi), plasma glucose andcatecholamine levels. In series III, additional measurements were made of whole-blood oxygen tension (PaO2), content (Cao2) and plasma lactate ([lactate]). pHe,PaO2 and Cao2 were determined as described by Vermette & Perry (1988). Plasmawas separated by centrifugation (2min at 13000g), acidified (0-6 moll"1 HC1O4),and stored (—20°C) for later determination of plasma glucose levels. Plasmasamples (100-200^1) were also stored for subsequent determination of catechol-amines and [lactate] (series III only) at — 70 °C. Plasma glucose and lactate levelswere measured enzymatically according to Bergmeyer (1974). Epinephrine andnorepinephrine levels were measured by HPLC and electronic detection, asdescribed in detail by Woodward (1982).

Liver and muscle glucose and glycogen concentrations were determined infollowing manner. Frozen tissue samples were weighed, deproteinized in ice


HCIO4 (0-6moll"1, 1:4 dilution), and homogenized. A sample of neutralizedsupernatant was hydrolyzed with amyloglucosidase (in acetate buffer, pH 4-6-4-8)and glucose was determined as above. Glycogen (in /xmol glucosyl units g"1 tissue)was determined as the difference in glucose concentrations in the hydrolyzed andnonhydrolyzed samples.

The activities of GPase and PK were determined spectrophotometrically byrecording the change in absorbance of NAD(P) at 340nm (Emu = 6-22). Livertissue was sonicated (Koutes MicroUltrasonic cell disrupter) in aphosphorylation-dephosphorylation 'stopping buffer' (5:1 w/v; 50 mmol I"1

imidazole, 15 mmol I"1 /J-mercaptoethanol, 100 mmol I"1 KF, 5 mmol F 1 EDTA,5mmoll - 1 EGTA, pH7-5, at room temperature; after Stalmans & Hers, 1975),layered onto a 3 ml Sephadex (G-25) column equilibrated with stopping buffer,and centrifuged at low speeds according to Christopherson (1983) to remove smallmetabolites. PK activity was determined in SOmmoll"1 imidazole buffer (pH7-5)containing lOmmoll"1 MgCl2, 2-Smmoir1 ADP, lOOmmoll"1 KC1,0-12 mmol I"1 NADH, excess lactate dehydrogenase (10 units) and either 0-2, 0-4or 5-0 mmol I"1 phosphoenolpyruvate (PEP, omitted for control). GPase activitywas determined in SOmmoll"1 phosphate buffer (pH7-0) containing 0-4 mmol I"1

NADP+, 15 mmol I"1 MgSO4, 0-2 mmol I"1 glucose-l,6-bisphosphate, 2 mmol T 1

AMP, excess glucose-6-phosphate dehydrogenase and phosphoglycerate mutase(each lunit), 2mgml~1 dialyzed glycogen (omitted for control) and either0-25mmoir1 EDTA or lOmmolP1 EDTA and lOmmoll"1 caffeine. Caffeineinactivates the inactive or b form of GPase and therefore provides a direct measureof the active or a form of the enzyme. GPase a activity is defined as that in thepresence of caffeine and total GPase as activity without caffeine (Stalmans &Hers, 1975).

Cyclic AMP was measured in liver tissue using a cyclic-[3H]AMP kit (Amer-sham Corp.). Tissues were homogenized in 0-7% perchloric acid containingEDTA, neutralized and analyzed as directed.

Results are given as means ± 1 standard error (S.E.) of the mean. Initial andfinal blood sample values were compared using a Student's paired Mest (P < 0-05).Comparisons between groups of fish were performed using a Student's unpairedMest (P<0-05).

Results

Pyruvate kinase kinetics

The substrate-dependence of pyruvate kinase is shown in Fig. 1A. The substrateconcentration for half-maximal saturation (So-5) for the enzyme with respect tophosphenolpyruvate (PEP) was 0-60 mmol I"1 (Fig. IB) and Vm at 15°C was^ • l / z m o l g ^ m k r 1 (Fig. 1A). The Hill constant («H) . calculated from the slopeof the line in Fig. IB, was 1-80. An nH value greater than 1 indicates positivefcoperativity in the binding between ligand and regulatory enzyme. L-Alaninep mmol I"1) did not significantly modify the kinetic properties of PK with respect

174 P. A. WRIGHT, S. F. PERRY AND T. W. MOON

25T A 3 B

101

1 2 3 4[PEP] (mmoir1)

6-

4-

2-

0 J

0-00 0-02 0-04 0-06 0-08 010[F-1.6-BP] (mmoir1)

Hs

I OH

- 1

- 2 J

10 n

8-

6-

0-5

00

"5 4-

2-

D

F-1.6-BPIT~ " '

Control

1 2 3[Alanine] (mmoir1)

Fig. 1. (A) Phosphoenolpyruvate (PEP) saturation curves of pyruvate kinase in wholeliver from intact trout (15°C) with no modulators (solid line, N = 5), with the additionof 0-1 mmoir1 fructose-l,6-bisphosphate (F-1.6-BP) (dotted line, N=S) or2mmoir1 alanine (dashed line, N=6). (B) Hill plots of data in A. (C) F-1,6-BPsaturation curve of pyruvate kinase at O-Smmoll"1 PEP (10°C, N = 4). (D) Alaninesaturation curves of pyruvate kinase at 0-8mmoir1 PEP with no modulator (control,solid line) and with O-lmmoll"1 F-1.6-BP (dashed line) (10°C, N=3). Bars indicate±lS.E.M.

to PEP (S0.5 = 0-66mmoi - 1 Vm= 15 min"1, nH = l-84; Fig. 1A,B);this effect was independent of alanine concentration or the presence of theactivator fructose-l,6-bisphosphate (F-1,6-BP) (Fig. ID). In contrast, F-1.6-BP(O-lmmoll"1) had a strong activating effect, decreasing S0-5 from 0-53 to0-07mmoir1 (Fig. 1A,B). This stimulation of PK by F-1,6-BP occurs at lowactivator concentrations (Fig. 1C), with a Ka value of O-rjO/xmoll"1. F-1,6-BPdecreased the «H value from 1-80 to 0-99, indicating an absence of cooperativebetween ligand (PEP) and enzyme.


It is not possible to determine the S0.5 for GPase with respect to glycogenbecause of the presence of unknown, but substantial, levels of endogenous tissueglycogen.

Plasma catecholamine levels

It is clear that plasma catecholamine levels increased in response to epinephrineand norepinephrine injections and hypoxia (Fig. 2). Because of the large varia-bility in the magnitude of the responses, however, the changes were generally not

120-1 A Epinephrine

100

"o 60-c

40

20

0

100

60

40-

20

B Norepinephrine

C E E+P N N+P P HC H H+P C E E+P N N+P P HC H H+P

160-

140-

120

100-

i so-E

C 60-1

40

20-1

0

C Total catecholamines * *

C E E+P N N+P P HC H H+P

Fig. 2. Plasma epinephrine (A), norepinephrine (B) and total catecholamines(A+B = C) innmoll for control (C) series I and II, N= 11; epinephrine (£), N = 6;epinephrine + propranolol (E+P), N=7; norepinephrine (N), N = 6; norepinephrine+ propranolol (N+P), N=6; propranolol series I and II (P), N= 12; control series III(HC), N = 6; hypoxia (H), N=8; and hypoxia + propranolol (H+P), N = 6. Initiallevels in each group of fish are shown by the filled bars and final levels, following theexperimental treatment, by the open bars. Bars indicate ±1S.E.M. (*S.E. =201 and**S.E. =255).


significant. Control catecholamine levels were approximately lnmoll \ exceptfor fish pre-treated with propranolol, where initial levels were two- to sevenfoldhigher (Fig. 2A, 2B). With epinephrine and norepinephrine injections, the finalplasma concentration was 60 nmol I"1 in both cases. Hypoxia elevated epinephrine(22nmoll~1, Fig. 2A) and, to a smaller extent, norepinephrine levels (7 nmol I"1,Fig. 2B). Pre-treatment with propranolol prior to hypoxia resulted in a consider-ably larger total catecholamine release to the plasma (l^WrimolP1, Fig. 2C).

Series I: epinephrine

Epinephrine elevated plasma glucose levels (46%) relative to initial values(Table 2A). Liver glucose also increased compared with control values, but thiswas not significant. There were no changes in muscle glucose levels and, althoughmuscle glycogen levels decreased, high variability masked any significant trends.Pre-injection with propranolol eliminated the increase in glucose followingepinephrine treatment, implying that the epinephrine effect was mediated by/S-receptors. Propranolol alone had no significant effect on glucose, but liver and

Table 2. (A) Plasma glucose, liver and muscle glucose and glycogen and(B) blood pHe and pHi data for series I epinephrine experiments

Plasma glucose Liver MuscleInitial Final Glucose Glycogen Glucose Glycogen

ASaline control 3-32 ±0-60 3-20 ±0-48 2-49 ±0-23 47-31 ± 14-98 0-93 ±0-15 12-72 ±3-42

(N=6)Epinephrine 2-72 ±0-28 3-97 ±0-23* 4-40 ± 1-57 42-42 ±25-03 0-81 ± 0-04 7-90 ±1-11

(N=6)Propranolol control 2-94 ±0-30 3-09 ±0-20 2-62 ±0-20 108-37 ± 15-58 0-69 ± 0-04 49-12 ±2-22

(TV = 8)Propranolol-I- 3-46 ±0-37 3-47 ±0-26 3-64 ±0-57 64-10 ±17-16 0-89 ±0-19 11-72 ±1-30

epinephrine(N=8)

InitialpHe

Final InitialpHi

Final

BSaline control 7-84 ±0-03 7-90 ±0-05 7-41 ± 0-02 7-42 ±0-03Epinephrine 7-84 ±0-03 7-89 ±0-05 7-39 ±0-03 7-33 ±0-05Propranolol 7-80 ±0-03 7-79 ±0-02 7-47 ±0-05 7-41 ±004Propranolol + epinephrine 7-81 ±003 7-80 ±004 7-47 ± 0-03 7-37 ±0-03

Initial samples were taken 30 min after the pre-injection (saline/propranolol) and final samples weretaken 15 min after the experimental injection (saline/epinephrine).

Units for glucose are /tmolg"1 and for glycogen /anol glucosyl units g"1 liver or muscle tissue.Means ± IS.E. (N= number of animals).* Significantly different from pre-injection value, paired Mest, /><0-05.


Table 3. (A) Liver pyruvate kinase activity for series I at 0-2 and 5-0mmoll 'phosphoenolpyruvate (PEP) concentrations and the ratio of activities at0-2/5-OPEP and (B) glycogen phosphorylase activity presented as totalGPase fa + b), GPase a, and the relative percentage of the active form to the total

activity [percent GPase a = a/fa + b)]

ASaline control

(W=6)Epinephrine

(TV =6)Propranolol control

(TV =8)Propranolol + epinephrine

BSaline controlEpinephrinePropranolol controlPropranolol + epinephrine

0-2

0-68 ±0-12

0-28 ±0-04*

l-30±0-29t

l-06±0-21t

Total GPase

1-27 ±0-331-36 ±0-100-40±0-08*'t0-47±0-10*'t

PEPOnmolP1)5-0

9-23 ±1-06

10-63 ±1-09

12-21 ±1-23

15-19 ±2-13

GPase a

0-50 ±0190-69 ±0-140-10±0-03*-t0-13±0-05t

0-2/5-0

007 ±001

003 ± 0-00*

0-12±0-03t

0-08 ± 0-02t

Percent GPase a

34±749 ±723±5f

24±5t

Activities in/imol min lg l at 15°C.Means ± ls.E.•Significantly different from control, unpaired /-test, P<0-05.t Significantly different from epinephrine, unpaired /-test, P<0-05.

muscle glycogen levels were well above values in the other treatments. pHe andpHi were not altered by any of the experimental treatments (Table 2B).

Epinephrine significantly modified PK activity (Table 3A). At low PEP concen-trations (O-ZmmolP1), activity was reduced two- to fivefold compared with thecontrol and the other treatments. The ratio of activity rates measured at low(O-Immoll"1) and saturating (SmmolP1) concentrations of PEP provides anindication of the phosphorylation state of PK: that is, a decrease in the ratiocorresponds to an increase in PK phosphorylation. The ratio of activities(Table 3A), was significantly lower in fish treated with epinephrine compared withvalues for saline and both propranolol treatments; as maximal activities(SmmolP1 PEP) remained unchanged, this decrease indicates an increase in theSo-s (PEP) value. Propranolol alone increased the ratio and eliminated theinhibitory effect of epinephrine on PK.

In preliminary experiments, PK activities were estimated in the presence of^ o l l " 1 L-alanine and Olmmoll" 1 F-1,6-BP under all experimental treat-ments. F-1,6-BP had a strong stimulatory effect on PK activity, increasing the


0-2/5-0PEP ratio; however, this effect was independent of experimental treat-ment. Alanine did not significantly modify PK activity in any of the experimentalmanipulations. These data are not shown since they do not add any newinformation to that presented in Fig. 1.

GPase values in epinephrine-treated fish were not significantly different fromcontrol values. The active form of GPase (percent GPase a), total GPase (a+b)and GPase a activities, however, were significantly higher in epinephrine-treatedfish than in the propranolol control and propranolol + epinephrine experiments(Table 3B). Propranolol alone significantly reduced GPase activity (total and a)compared with the saline control value.

Series II: norepinephrine

The results of the norepinephrine experiments (Tables 4, 5) were similar tothose of series I for epinephrine. The major difference was that at the samecirculating concentration as epinephrine ((iOnmoll"1, Fig. 2B), the effects ofnorepinephrine on the measured variables were smaller. Following norepineph-rine injection, plasma glucose level increased significantly (16 % ) , but this increase

Table 4. (A) Plasma glucose, liver and muscle glucose and glycogen and (B) blood pHeand pHi data for series II norepinephrine experiments


ASaline control 3-93 ±0-46 3-76 ±0-32 2-65 ±0-24 115-59 ±31-62 0-72 ±0-12 8-22 ±1-53

(AT = 6)Norepinephrine 3-64 ±0-22 4-25 ± 0-40* 3-28 ± 0-47 62-89 ±13-56 0-91 ±0-13 8-84 ±1-29

(TV =7)Propranolol control 4-02 ±0-39 3-89 ±0-42 2-73 ±0-07 132-23 ±40-87 0-95 ±0-22 11-46 ±2-72

(TV = 5)Propranolol + 2-83 ±0-54 2-88 ±0-43 2-58 ±0-50 80-63 ±33-57 0-87 + 0-25 11-17 ±2-68

norepinephrine(TV =6)

pHe pHiInitial Final Initial Final

BSaline control 7-86 ±002 7-88 ±0-02 7-22 ±005 7-26 ±0-01Norepinephrine 7-91 ± 0-02 7-89 ±0-04 719 ±0-03 7-22 ±004Propranolol 7-77 ±0-03 7-75 ±0-02 7-31 ±0-06 7-19 ±0-10Propranolol + norepinephrine 7-81 ±003 7-81 ±0-05 7-36 ±0-07 7-39 ±008

Initial samples were taken 30min after the pre-injection (saline/propranolol) and final samples weretaken 15min after the experimental injection (saline/norepinephrine).

Units for glucose are /imol g~l and for glycogen janol glucosyl units g~1 liver or muscle tissue.Means ± 1 S.E. (TV = number of animals).•Significantly different from pre-injection value, paired Mest, P<0-05.


Table 5. (A) Liver pyruvate kinase activity for series II at 0-2 and 5-0mmoll 'phosphoenolpyruvate (PEP) concentrations and the ratio of activities at0-2/5-0 PEP and (B) glycogen phosphorylase activity presented as totalGPase fa + b), GPase a, and the relative percentage of the active form to the total


ASaline control

(JV=6)Norepinephrine

(N = l)Propranolol control

(TV = 5)Propranolol + norepinephrine

(TV = 8)

BSaline controlNorepinephrinePropranolol controlPropranolol + norepinephrine

0-2

0-64 ±0-26

0-39 ±0-07

1-08 ±0-58

0-76 ±0-25

Total GPase

0-75 ±0-201-24 ±0-380-48 ±0-130-70 ± 0-27

PEPfjnmoir1)5-0

10-26 ±1-07

11-49 ±1-64

8-62 ± 1-22

10-20 ±1-18

GPase a

0-17 ±0-060-53 ±0-230-13 ±0-060-28 ±0-15

0-2/5-0

0-10 ±0-03

0-03 ± 0-01*

0-13 ±0-05

007 ± 0-02

Percent GPase a

19 ± 430±724 ± 429 ± 7

Activities in/unol min xg ' at 18°C.Means ± IS .E .* Significantly different from control, unpaired /-test, P<0-05.

was only half of that observed with epinephrine (Table 2). Pre-injection withpropranolol eliminated the increase in glucose associated with norepinephrinetreatment, again suggesting mediation by /5-receptors. pHe and pHi were notaltered by any of the experimental treatments (Table 4B).

PK activity at O^mmoll"1 PEP was reduced by norepinephrine compared tothe other treatments, but this effect was not significant. Norepinephrine, however,did significantly depress the 0-2/5-0 PEP ratio compared with the saline controlvalue.

Norepinephrine increased total GPase, GPase a and percent GPase a activities(Table 5A), but these trends were not significant.

Series III: hypoxia

Plasma epinephrine and norepinephrine were both elevated after 30 min ofhypoxia, although epinephrine levels were three times those of norepinephrinekig. 2). Plasma glucose levels were constant during hypoxia. The combination ofpropranolol and hypoxia, however, caused a significant reduction of plasma


glucose (20%) compared to pre-treatment levels. There were no significantchanges in liver or muscle glucose and glycogen levels. Red cell pHi wassignificantly elevated during hypoxia, which indicates /S-adrenoreceptor stimu-lation (Tetens & Christensen, 1987). Blood P a ^ and Ca^ were severely depressedduring hypoxia. Pre-treatment with propranolol significantly reduced initial bloodoxygen levels, and hypoxia compounded this effect by lowering PaOl and Ca^values below those seen with hypoxia alone (Table 6C). Hypoxia resulted in athree- to fourfold increase in plasma [lactate], indicating an activation of anaerobicmetabolic pathways.

PK activity remained-constant during hypoxia (Table 7). Pre-treatment withpropranolol plus hypoxia, however, caused a significant increase in PK activity atO^mmoll"1 PEP, and an increase in the 0-2/5-0PEP ratio (Table 7). Propranololalone decreased maximal PK activity (at 5-Ommoll"1 PEP), although a similarchange at O ^ m m o i r 1 PEP meant that there was no change in the 0-2/5-OPEPratio.

Hypoxia increased total GPase activity by two- to threefold over saline andpropranolol control treatments. GPase a activity was 5-7 times greater duringhypoxia compared to values for propranolol treatment with or without hypoxia.Percent GPase a activity was significantly higher during hypoxia compared withvalues during treatments with propranolol + hypoxia (Table 7).

Cyclic AMP

Liver cyclic AMP levels increased slightly with epinephrine, norepinephrine andhypoxia, but none of these treatments resulted in significant changes comparedwith control levels (Table 8).

Discussion

Liver pyruvate kinase and glycogen phosphorylase activities in vivo

Pyruvate kinase in teleost liver tissue was initially reported to be nonregulatory(Somero & Hochachka, 1968; Guderley etal. 1978; Moon & Hulbert, 1980;Guderley & Cardenas, 1980). The results of this study, however, and those of fourrecent investigations (Petersen etal. 1988; Rahman & Storey, 1988; Sand, 1988;Moon et al. 1989) indicate the contrary. An explanation for this discrepancy is thatdifferences in the physiological state of the fish and/or the preparation of the liverhomogenate result in variable states of phosphorylation of the enzyme. Thesensitivity of PK to allostenc modulators is dependent on whether the enzyme isphosphorylated (inactive state) or dephosphorylated (active state) (Pilkis et al.1988). Furthermore, the procedure for isolating hepatocytes in rats, which issimilar to that in trout, results in the progressive dephosphorylation of PK (Riouetal. 1985). The substrate concentration required for half-maximal saturation(So 5) is dependent on the phosphorylation state, and this may explain why ourin vivo So 5 value (O^mmolP1 PEP) is an order of magnitude greater than thtfmeasured by Petersen et al. (1988) in trout hepatocytes. Hence, kinetic parameter


Table 6. (A) Plasma glucose, liver and muscle glucose and glycogen, (B) pHe and pHiand (C) PaO2, CaO2 and plasma lactate data for series III hypoxia experiments


ASaline control 3-12 ±0-95 3-37 ±1-03 2-28 ±0-66 64-98 ±25-07 0-60 ±0-17 8-67 ±1-97

(N = 8)Hypoxia 5-72 ±1-69 6-12 ±1-75 4-36 ± 1-08 105-13 ±32-16 0-76 ±0-18 8-58 ±2-00

( t f= l l )Propranolol control 4-02 ±0-39 3-89 ±0-42 2-73 ±0-07 132-23 ±40-87 0-95 ±0-22 11-46 ±2-72

Propranolol +hypoxia(N=6)

4-45±0-87 3-54 ± 0-65* 2-50 ± 0-40 139-00 ±33-72 0-69 ±0-18 10-48 ±1-60

InitialpHe

Final InitialpHi

Final

BSaline controlHypoxiaPropranolol controlPropranolol + hypoxia

7-91 ±0-047-83 ±0-037-77 ±0-037-78 ±0-02

7-94 ±0-027-76 ±0-057-75 ±0-027-68 ±0-10

7-38 ±0-047-27 ±0-027-31 ±0067-22 ±0-03

7-35 ± 0-0417-46 ±0-03*7-19 ±0-10t7-23±0-08t

Initial(kPa)

Final(vols%)

Initial Final[Lactate] (janol 1 x

Initial Final

Saline control 16-0 ±1-3 18-3 ±0-9 7-37 ±0-72 6-67 ±0-47 1-13 ±0-35 1-28 ±0-31Hypoxia 15-5 ±1-2 5-9±l-5+ 8-38 ±0-62 3-84 ±0-29* 1-16 ±0-20 4-47 ±0-67*'**Propranolol + 9-3 ±0-003"'t 3-3 ±0-01* 6-50 ± 0-03t 2-04 ±0-08* 0-96 ±0-30 2-66 ±0-76*

hypoxia

Initial samples were taken 30min after the pre-injection (saline/propranolol) and final samples30min after the initiation of external hypoxia.

Units for glucose are /zmolg"1 and for glycogen jimol glucosyl unitsg"1 liver or muscle tissue.Means ± Is.E. (N = number of animals).•Significantly different from initial value, paired Mest, P<0-05.••Significantly different from control value, unpaired Mest, P<005 .t Significantly different from hypoxia value, unpaired Mest, P<0-05.JData from series II.

for PK measured in thus study, using freeze-clamped liver tissue and aphosphorylation-dephosphorylation stopping buffer, may be closer to in situvalues than those measured on isolated hepatocytes or in many of the previousstudies made on this enzyme in fish.^Alanine had virtually no effect on PK activity in the present study, but is a stronginhibitor of mammalian liver and invertebrate hepatopancreas PK (see Munday


Table 7. (A) Pyruvate kinase activity for series III at 0-2 and 5-0mmoll 1

phosphoenolpyruvate (PEP) concentrations and the ratio of activities at0-215-0 PEP and (B) glycogen phosphorylase activity presented as totalGPase fa + b), GPase a, and the relative percentage of the active form to the total


ASaline control

Hypoxia(JV=11)

Propranolol control

Propranolol + hypoxia(TV = 6)

0-2

1-3010-31

1-1310-23

1-0810-58

2-36!0-57f

Total GPase

PEPfmmoir1)5-0

13-1810-82

13-2311-45

8-62 11-22*

10-0211-42

GPase a

0-2/5-0

0-1110-03

0-0910-02

0-1310-05

0-24 + 0-04*'t

Percent GPase a

B

Saline controlHypoxiaPropranolol control^Propranolol + hypoxia

0-7710-161-3910-18*0-4810-13t0-3610-llt

0-2210-080-5210-100-13±0-06t0-07!0-03t

23143414241413±6t

Activities in janol min lg x at 16°C.Means! ls.E.* Significantly different from control, unpaired Mest, P<0-05.t Significantly different from hypoxia, unpaired Mest, P<0-05.$ Data from series n .

etal. 1980). Moon etal. (1989) have also shown an effect of alanine on rainbowtrout liver PK; the reason for this discrepancy is not understood. In contrast,relatively low concentrations of F-1,6-BP activated PK, lowered the S0-5 value forPEP and modified the shape of the PEP saturation curve to a hyperbola. Thesensitivity of trout PK to F-1,6-BP activation has been reported previously inrainbow trout (Moon et al. 1989), goldfish (Rahman & Storey, 1988) and flounder(Sand, 1988).

The net rate of glycogen breakdown depends largely on the relative activity ofthe active form of GPase or percent GPase a. Glycogenolysis is stimulated duringstress in many vertebrates (Hems & Whitton, 1980) and, therefore, one wouldexpect activity of the a form to be low in resting animals and increase with stress.Published liver percent GPase a values for resting fish are high, ranging from 60 to90% (Ottolenghi etal. 1985, 1986; Perry etal. 1988; Mommsen etal. 1988; Moonetal. 1989), suggesting near maximal rates of glycogen breakdown under reconditions. These high percent GPase a values may possibly be an artifact of t


Table 8. Liver cyclic AMP levels

Control(#=19)

Propranolol(#=13)

Epinephrine(#=5)

Epinephrine + propranolol(#=8)

Norepinephrine(#=7)

Norepinephrine + propranolol(#=6)

Hypoxia(#=10)

Hypoxia + propranolol(#=5)

[cyclic AMP](pmol g"1 liver tissue)

538 ±29

562 ±48

644 ±81

484±56

601 ±34

510 ±44

580 ±63

468±72

Data from control and propranolol experiments were not significantly different and thereforewere pooled.

Means ± ls.E.

assay technique. Stalmans & Hers (1975) reported that AMP activated both the aand b forms of GPase and only when caffeine was added to liver homogenates wasthe b form completely inhibited. Their technique provided a more accurateestimate of percent GPase a. This study is one of the first to use the criteria ofStalman & Hers (1975), and reports percent GPase a values of between 10 and50 %, depending upon conditions. These values, to our knowledge, are the lowestreported for any teleost fish, including a recent study on rainbow trout (Moonet al. 1989), or for mammalian liver (e.g. 70-85 %; Stalmans & Hers, 1975; Hems& Whitton, 1980).

It should be noted that total GPase (a+b) was not constant between differentexperimental treatments (Tables 3B, 5B, 7B). Although this observation has beenreported by others (Ottolenghi et al. 1986,1988; Mommsen et al. 1988; Perry et al.1988), we do not have an explanation for these results.

Catecholamine regulation of glucose metabolism during hypoxia

The results of this study support the hypothesis that catecholamines, releasedduring hypoxia, have an important role in regulating the availability of glucose.Injections of epinephrine or norepinephrine into resting fish increased plasmaglucose levels, and this hyperglycemia was blocked by pre-treatment with theyS-adrenoreceptor antagonist propranolol. During hypoxia, plasma catecholamine

tls increased but there was no change in plasma glucose levels. We suggest thatreason we did not observe catecholamine-mediated hyperglycemia was


because there was an increase in tissue glucose utilization, as a result of inhibitionof aerobic, and activation of anaerobic, metabolism (Dunn & Hochachka, 1987;Boutilier etal. 1988). Plasma lactate levels increased fourfold during hypoxia,supporting an enhancement in anaerobic metabolism. Finally, plasma glucoselevels were significantly depressed during hypoxia after blockade of/S-adrenoreceptors. This indicates an essential role for /5-adrenoreceptor-mediatedmobilization of glucose during hypoxia to meet metabolic requirements.

Catecholamines regulate glucose metabolism, in part by modifying the activityof two key regulatory enzymes: PK, a regulatory enzyme in glycolysis andgluconeogenesis, and GPase, the enzyme directly responsible for glycogenbreakdown. Injections of epinephrine, and to a lesser extent norepinephrine,increased the percent GPase a (statistically significant compared with propranolol-treated fish) and decreased PK activity in the liver, indicating an activation ofglycogenolysis and an inhibition of glycolysis and/or stimulation of gluconeogen-esis, respectively. The catecholamine effects on GPase and PK were blockedcompletely by propranolol, identifying /J-receptors as the mediators of the trouthepatic catecholamine response. Blockade of ^-adrenoreceptors caused substan-tial reductions of percent GPase a and total GPase activities, while PK activity wasmarkedly stimulated. Thus, it is apparent that the maintenance of blood glucoselevels during hypoxia was due to adrenergic activation of GPase and inhibition ofPK. In turn, these enzymatic changes accelerated hepatic glycogenolysis andgluconeogenesis while inhibiting glycolysis. The unusually high activity of PK(assayed at low PEP concentrations) and low activity of GPase during hypoxiaafter /J-adrenoreceptor blockade suggest that internal hypoxia is specificallyaffecting the phosphorylation state, that is dephosphorylating these two enzymes.Thus, it would appear that the mobilization of catecholamines during hypoxiaserves to offset the deleterious effects of hypoxemia on PK and GPase.

The hyperglycemic effect of catecholamines and involvement of/J-adrenoreceptors have been demonstrated in vitro in trout (Mommsen et al.1988) and in catfish (Brighenti et al. 19876; Janssens & Lowrey, 1987). The presentstudy is the first to show ^-adrenoreceptor-mediated catecholamine effects onglucose metabolism in the intact trout with administration of physiologicallyrelevant epinephrine or norepinephrine levels. In addition, we have shown thatliver percent GPase a is twofold greater in epinephrine-treated fish than inpropranolol- or propranolol + epinephrine-treated fish, whereas previous hepato-cyte studies have failed to demonstrate a marked change (Ottolenghi et al. 1985,1986; Mommsen et al. 1988). Epinephrine inhibits PK activity in trout hepatocytes(Mommsen et al. 1988), although the magnitude of the effect was considerably lessthan we report in the present study. Mommsen and co-workers found that the0-1/5 0 PEP ratio decreased by 20%, compared to our results in which the0-2/5-0PEP ratio was reduced by 60%. As discussed above, the hepatocyteisolation procedure may have altered the phosphorylation state of PK, resulting ina reduced sensitivity to catecholamines. Norepinephrine mimicked the effeepinephrine but was less potent (Tables 4, 5). Similar results were obtained


Brighenti et al. (1987a,£>) in catfish hepatocytes. In trout hepatocytes, however,both hormones induced similar changes in glucose production and lactategluconeogenesis (Mommsen et al. 1988).

In many animals, the immediate response to oxygen deprivation is thebreakdown of liver glycogen and subsequent mobilization of glucose via the bloodto other tissues for anaerobic utilization. In the longer term, as glycogen levelsdecline, liver gluconeogenesis must be activated to supply glucose required foranaerobic metabolism or, alternatively, overall metabolism may be depressed.Our results did not show a significant drop in liver or muscle glycogen storesfollowing 30min of hypoxia (Table 6A), but the increases in liver total GPase andpercent GPase a activity (Table 7B) suggest that glycogen catabolism may havebeen enhanced. Liver glycogen levels are extremely variable in fish and withoutinitial glycogen contents it is difficult to evaluate the glycogenolytic effect ofhypoxia. In fish hepatocytes and perfused livers, glycogen level decreases in thepresence of catecholamines (Janssens et al. 1983; Ottolenghi et al. 1985, 1986;Brighenti et al. 1987a; Janssens & Lowery, 1987). Dunn & Hochachka (1987)found that trout exposed to water with a P Q , of 4 kPa for 3 h showed no significantfall in liver glycogen. Thus, it may be that 30min of moderate hypoxia(Pwch ~ 7-1 kPa) is not of sufficient intensity and duration to cause a discernibledepletion of glycogen stores, despite elevated catecholamine levels.

Catecholamine release during hypoxia is dependent on the intensity andduration of the hypoxic stress (Tetens & Christensen, 1987; Boutilier et al. 1988).Plasma epinephrine levels in trout exposed to hypoxia (water PWQJ = 7-1 kPa for30min, Fig. 2A) were two- to 10-fold greater than that reported by Fievet et al.(1987) and Tetens & Christensen (1987) for trout in water of AvO2 = 5-3 kPa for20-30 min. Norepinephrine levels (Fig. 2B) were lower than in the other twostudies (Fievet et al. 1987; Tetens & Christensen, 1987) and, therefore, it may bemore instructive to look at the total catecholamine levels ([epinephrine + norepi-nephiine]; Fig. 2C). Total plasma catecholamine concentration was 29nmoll~1

during hypoxia (Fig. 2C), a value very similar to the total level in the Fievet et al.(1987) study and 2-5-fold higher than levels reported by Tetens & Christensen(1987). Control or initial catecholamine levels were approximately lnmoll"1 ,which is representative of true resting values (Mazeaud & Mazeaud, 1985). Stresslevels of epinephrine and norepinephrine may be as high as 200-400 nmol I"1 and,therefore, the measured levels of either catecholamine (60 nmol I"1) after injectionin the present study were well within the physiological range, although higher thanthose achieved during hypoxia. An interesting observation is that propranololalone stimulated catecholamine release and, during hypoxia, resulted in a higherplasma catecholamine concentration (Fig. 2). Tetens & Christensen (1987) alsonoted a catecholamine stimulation with propranolol, although they did notcomment on its significance. The fact that pre-treatment with propranolol reducedblood PaOl and CaO2 (Table 6C) suggests that fish were hypoxic before the

Rberimental treatment (epinephrine, norepinephrine or hypoxia). This mightplain the initial small rise in both epinephrine and norepinephrine in fish treated


with propranolol. The marked increase in catecholamines with both propranololand hypoxia, compared to hypoxia alone, is presumably related to the muchgreater hypoxic stress (PaO2 = 3-3 kPa, CaOz = 2 vols%, Table 6C) in these fish.

Resting or baseline catecholamine levels ( lnmolP1) may be important in theregulation of glucose metabolism. When propranolol was administered to restingfish in series I, total GPase and GPase a activities were reduced by three- tofivefold compared to control values (Table 3B). (The same trends were observedin series II, but the differences were not statistically significant.) There are twopossibilities; catecholamines tonically regulate glycogenolysis in the liver, or pre-mortem handling induced a catecholamine surge that resulted in an immediate/3-adrenoreceptor response (Moon etal. 1988). In this study, the procedure ofkilling the fish, excising the liver and freezing the tissue in liquid N2 took less than10s. It is possible, however, that some catecholamines were released within thisshort time. To differentiate between tonic adrenergic control of GPase or pre-mortem stress, further studies on intact fish are necessary where the potentialproblem of stress-related handling is reduced or eliminated.

Binding of catecholamines to the /S-adrenoreceptor activates adenylate cyclaseand increases cellular cyclic AMP levels (see Levitzki, 1988). In the present study,cyclic AMP levels did increase slightly with catecholamines and hypoxia, althoughthe changes were not significant. In contrast, cyclic AMP levels increased rapidlywith both epinephrine or norepinephrine treatment in fish hepatocyte studies(Janssens & Lowrey, 1987; Brighenti etal. 1987a). There are several possibleexplanations for the discrepancy between in vivo and in vitro results. First,stimulation of cellular cyclic AMP levels is dose-dependent. A plasma catechol-amine level in vivo, however, is probably not physiologically equivalent to thesame in vitro dose, because of the more complex structure and circulation of thewhole organ versus isolated hepatocytes. Second, the sampling time of our in vivostudy may not have corresponded with the maximum surge of intracellular cyclicAMP levels. Based on the ubiquitous nature of cyclic AMP as the /S-adrenoceptorintracellular messenger in higher vertebrates (Levitzki, 1988), teleosts (Janssens &Lowrey, 1987; Brighenti etal 1987a) and amphibians (Janssens etal. 1986), it isprobable that the low (but physiological) catecholamine dose used in our study,the inherent variability of in vivo data and transient surges have masked the role ofcyclic AMP as the intracellular messenger in trout liver tissue.

We thank Dr Tom Mommsen for his helpful discussions during the initial stagesof this study. This work was supported by NSERC of Canada operating andequipment grants to TWM and SFP. PAW was the recipient of a KillamPostdoctoral Fellowship.

ReferencesBERGMEYER, H. U. (1974). Methods of Enzymatic Analysis. New York: Academic Press.BIRNBAUM, M. J., SCHULTZ, J. & FAJN, J. N. (1976). Hormone-stimulated glycogenolysij

isolated goldfish hepatocytes. Am. J. Physiol. 231, 191-197.


BOUTIUER, R. G., DOBSON, G., HOEGER, U. & RANDALL, D. J. (1988). Acute exposure tograded levels of hypoxia in rainbow trout Salmo gairdneri: metabolic and respiratoryadaptations. Respir. Physiol. 71, 69-82.

BRJGHENTI, L., PUVIANI, A. C , GAVIOLI, M. E., FABBRI, E. & OTTOLENGHI, C. (1987a).Catecholamine effect on cyclic adenosine 3':5'-monophosphate level in isolated catfishhepatocytes. Gen. comp. Endocr. 68, 216-223.

BRIGHENTI, L., PUVIANI, A. C , GAVIOLI, M. E. & OTTOLENGHI, C. (1987b). Mechanismsinvolved in catecholamine effect on glycogenolysis in catfish hepatocytes. Gen. comp.Endocr. 66, 306-313.

BUTLER, P. J., TAYLOR, E. W. & DAVTSON, W. (1979). The effect of long term, moderate hypoxiaon acid-base balance, plasma catecholamines and possible anaerobic end products in theunrestrained dogfish Scyliorhinus canicula. J. comp. Physiol. 132, 297-303.

CHRISTOPHERSON, R. I. (1983). Desalting protein solutions in a centrifuge column. MethodsEnzymol. 91, 278-281.

DEROOS, R. & DEROOS, C. C. (1978). Elevation of plasma glucose levels by catecholamines inelasmobranch fish. Gen. comp. Endocr. 34, 447-452.

DUNN, J. F. & HOCHACHKA, P. W. (1987). Turnover rates of glucose and lactate in rainbow troutduring acute hypoxia. Can. J. Zool. 65,1144-1148.

FIEVET, B., MOTAIS, R. & THOMAS, S. (1987). Role of adrenergic-dependent H+ release from redcells in acidosis induced by hypoxia in trout. Am. J. Physiol. 252, R269-R275.

GUDERLEY, H. & CARDENAS, J. M. (1980). A study of the catalytic properties of pyruvate kinaseisozymes from salmon and an examination of their functional relationships. /. exp. Zool. 212,269-277.

GUDERLEY, H., FIELDS, J. H. A., CARDENAS, J. & HOCHACHKA, P. W. (1978). Pyruvate kinasefrom the liver and kidney of Arapaima gigas. Can. J. Zool. 56, 852-859.

HEMS, D. A. & WHITTON, P. A. (1980). Control of hepatic glycogenolysis. Physiol. Rev. 60,1-50.

JANSSENS, P. A., CAINE, A. G. & DIXON, J. E. (1983). Hormonal control of glycogenolysis andthe mechanism of action of adrenaline in amphibian liver in vitro. Gen. comp. Endocr. 49,477-484.

JANSSENS, P. A., KLEINEKE, J. & CAINE, A. G. (1986). Calcium-independent stimulation ofglycogenolysis by arginine vasotocin and catecholamines in liver of the axolotl (Ambystomamexicanum) in vitro. J. Endocrinol. 109, 75-84.

JANSSENS, P. A. & LOWREY, P. (1987). Hormonal regulation of hepatic glycogenolysis in thecarp, Cyprinus carpio. Am. J. Physiol. 252, R653-R660.

LEVITZKI, A. (1988). From epinephrine to cyclic AMP. Science 241, 800-805.MAZEAUD, M. M. & MAZEAUD, F. (1985). Adrenergic responses to stress in fish. In Stress in Fish

(ed. A. D. Pickering), pp. 49-55. New York: Academic Press.MOMMSEN, T. P., WALSH, P. J., PERRY, S. F. & MOON, T. W. (1988). Interactive effects of

catecholamines and hypercapnia on glucose production in isolated trout hepatocytes. Gen.comp. Endocr. 70, 63-73.

MOON, T. W., FOSTER, G. D. & PUSETSHAYA, E. M. (1989). Changes in peptide hormones andliver enzymes in the rainbow trout deprived of food for six weeks. Can. J. Zool. (in press).

MOON, T. W. & HULBERT, W. C. (1980). Regulatory properties of pyruvate kinases isolatedfrom tissues of the American eel, Anguilla rostrata le sueur. Comp. Biochem. Physiol. 65B,291-2%.

MOON, T. W., WALSH, P. J., PERRY, S. F. & MOMMSEN, T. P. (1988). Effects of in vivo beta-adrenoceptor blockade on hepatic carbohydrate metabolism in rainbow trout. Exp. Zool.248, 88-93.

MORATA, P., VARGAS, A. M., PITA, M. L. & SANCHEZ-MEDINA, F. (1982). Hormonal effects onthe liver glucose metabolism in rainbow trout {Salmo gairdneri). Comp. Biochem. Physiol.72B, 543-545.

VIUNDAY, K. A., GILES, I. G. & POAT, P. C. (1980). Review of the comparative biochemistry ofpyruvate kinase. Comp. Biochem. Physiol. 67B, 403-411.

NO, T. & TOMLINSON, T. (1967). Catecholamine and carbohydrate concentrations inainbow trout {Salmo gairdneri) in relation to physical disturbance. /. Fish Res. Bd Can. 24,

1702-1715.


OTTOLENGHI, C , PUVIANI, A. C , BARUFFALDI, A. & BRIGHENTI, L. (1984). Epinephrine effectson carbohydrate metabolism in catfish {lctalurus melas). Gen. comp. Endocr. 55, 378-386.

OTTOLENGHI, C , PUVIANI, A. C , BARUFFALDI, A., GAVIOLI, M. E. & BRIGHENTI, L. (1988).Glucagon control of glycogenolysis in catfish tissue. Comp. Biochem. Physiol. 90B, 285-290.

OTTOLENGHI, C , PUVIANI, A. C , GAVIOLI, M. E. & BRIGHENTI, L. (1985). Epinephrine effect oncarbohydrate metabolism in isolated and perfused catfish liver. Gen. comp. Endocr. 59,219-229.

OTTOLENGHI, C , PUVIANI, A. C , GAVIOU, M. E. & BRIGHENTI, L. (1986). Epinephrine effect onglycogen phosphorylase activity in catfish liver and muscle. Gen. comp. Physiol. 61, 469-475.

PERRY, S. F. & VERMETTE, M. G. (1987). The effects of prolonged epinephrine infusion on thephysiology of the rainbow trout, Salmo gairdneri. I. Blood respiratory, ionic and acid-basestates. /. exp. Biol. 128, 235-253.

PERRY, S. F., WALSH, P. L, MOMMSEN, T. P. & MOON, T. W. (1988). Metabolic consequences ofhypercapnia in the rainbow trout, Salmo gairdneri: /J-adrenergic effects. Gen. comp. Endocr.69, 439-447.

PETERSEN, T. D. P., HOCHACHKA, P. W. & SUAREZ, R. K. (1988). Hormonal control ofgluconeogenesis in rainbow trout: regulatory role of pyruvate kinase. J. exp. Zool. 243,173-180.

PILKIS, S. J., RAAFATEL-MAGHRABI, M. & CLAUS,T. M. (1988). Hormonal regulation of hepaticgluconeogenesis and glycolysis. A. Rev. Biochem. 57, 755-783.

RAHMAN, M. S. & STOREY, K. B. (1988). Role of covalent modification in the control ofglycolytic enzymes in response to environmental anoxia in goldfish. J. comp. Physiol. 157B,813-820.

Riou, J. P., AUDIGIER, C , LAVILLE, M., BEYLOT, M., PIGEON, P. & MORNEX, R. (1985).Dephosphorylation of L-pyruvate kinase during rat liver hepatocyte isolation. ArchsBiochem. Biophys. 236, 321-327.

RISTORI, M. T. & LAURENT, P. (1985). Plasma catecholamines and glucose during moderateexercise in the trout: Comparison with bursts of violent activity. Exp. Biol. 44, 247-253.

RUSH, S. B. & UMMINGER, B. L. (1978). Elimination of stress-induced changes in carbohydratemetabolism of goldfish (Carassius auratus) by training. Comp. Biochem. Physiol. 60A, 69-73.

SAND, O. (1988). The effect of fructose l,6diphosphate on pyruvate kinase from the liver of theflounder {Platichthys flesus L.). Comp. Biochem. Physiol. 90B, 401-407.

SOIVIO, A., WESTMAN, K. & NYHOLM, K. (1972). Improved method of dorsal aortacatheterization: haematological effects followed for three weeks in rainbow trout (Salmogairdneri). Finn. Fish. Res. 1, 11-21.

SOMERO, G. N. & HOCHACHKA, P. W. (1968). The effect of temperature on catalytic andregulatory functions of pyruvate kinases of rainbow trout and the antarctic fish Trematomusbernacchii. Biochem. J. 110, 395-400.

STALMANS, W. & HERS, H. G. (1975). The stimulation of liver phosphorylase b by AMP,fluoride and sulfate. Eur. J. Biochem. 54, 341-350.

TETENS, V. & CHRISTENSEN, N. J. (1987). Beta-adrenergic control of blood oxygen affinity inacutely hypoxia exposed rainbow trout. J. comp. Physiol. 157B, 667-675.

THORPE, A. & INCE, B. W. (1974). The effect of pancreatic hormones, catecholamines andglucose loading on blood metabolites in northern pike (Exos lucius L.). Gen. comp. Endocr.23, 29-44.

VERMETTE, M. G. & PERRY, S. F. (1988). Adrenergic involvement in blood oxygen transport andacid-base balance during hypercapnic acidosis in the rainbow trout, Salmo gairdneri. J. comp.Physiol. B 158, 107-115.

WOLF, K. (1963). Physiological salines for fresh water teleosts. Progve Fish Cult. 25, 135-140.WOODWARD, J. J. (1982). Plasma catecholamines in resting rainbow trout, Salmo gairdneri

Richardson by high pressure liquid chromatography. J. Fish Biol. 21, 429-432.WOSILAIT, W. D. & SUTHERLAND, E. W. (1956). The relationship of epinephrine and glucagon to

liver phosphorylase II enzymatic inactivation of liver phosphorylase. J. biol. Chem. 218,469-481.

Documents

REGULATION OF HEPATIC GLUCONEOGENESIS …hypoxia (with/without yS-adrenoreceptor blockade) on tissue glucose and glycogen levels, and on the kinetic behavior of two key enzymes involved